Optogenetic Induction of Pyroptosis, Necroptosis, and Apoptosis in Mammalian Cell Lines

Regulated cell death plays a key role in immunity, development, and homeostasis, but is also associated with a number of pathologies such as autoinflammatory and neurodegenerative diseases and cancer. However, despite the extensive mechanistic research of different cell death modalities, the direct comparison of different forms of cell death and their consequences on the cellular and tissue level remain poorly characterized. Comparative studies are hindered by the mechanistic and kinetic differences between cell death modalities, as well as the inability to selectively induce different cell death programs in an individual cell within cell populations or tissues. In this method, we present a protocol for rapid and specific optogenetic activation of three major types of programmed cell death: apoptosis, necroptosis, and pyroptosis, using light-induced forced oligomerization of their major effector proteins (caspases or kinases).


Background
Regulated cell death is a common feature of multicellular organisms and plays a key role in development and tissue homeostasis and in protecting the host against malignant growth and various pathogens. Research over the last two decades has identified over 12 different forms of regulated cell death (Galluzzi et al., 2018); however, their study is often complicated by the complex crosstalk and interconnectivity between the different cell death pathways (Bedoui et al., 2020). Additionally, the consequences of different types of cell death in the tissue still remain insufficiently understood, which is at least partially due to the challenges of selectively targeting single cells in multicellular populations, as well as to the pleiotropic effects of commonly used natural cell death triggers both on the dying cells and their neighbors. In recent years, multiple strategies have been developed to specifically ablate cells both in vitro and in living animals. However, some of these methods (such as laser ablation or photosensitization) still lack specificity regarding the type of cell death to be induced (Tirlapur et al., 2001;Qi et al., 2012), while others [such as chemically inducible dimerization of apoptotic or necroptotic effector proteins (Oberst et al., 2010;Wu et al., 2014)] suffer from a limited spatiotemporal control and require a delivery of soluble ligands, thus limiting their in vivo applications. To overcome these limitations and expand the scope of the tools available for programmed cell death induction, we recently developed a set of optogenetically activated cell death effectors (optoCDEs) (Shkarina et al., 2022), which enable selective induction of three major types of programmed cell death: apoptosis, pyroptosis, and necroptosis. These tools consist of three modules: 1) a photoactuator domain Cry2olig (Cry2 E490G), which responds to blue light by rapid homo-oligomerization, 2) an mCherry tag, which enables the detection of the cells expressing optoCDEs and estimation of the relative construct expression levels, and 3) an effector module ( Figure 1A-1C). For opto-caspases, the effector module corresponds to the protease domain (p20 and p10 subunits) of corresponding caspases; the endogenous linkers and cleavage sites essential for the caspase activation are retained, while CARD (in caspase-1, -4, -5, -9, and -11) and DED (caspase-8) domains, responsible for the endogenous upper-level protein-protein interactions and homo-oligomerization, are removed. In opto-RIPK3, the design is similar, while the RHIM motif (responsible for the upstream interaction with the RIPK1) is mutated. In optoMLKL, the effector domain orientation in relation to Cry2olig and mCherry is reversed to keep the MLKL N-terminus (responsible for membrane binding and disruption) exposed. The considerations behind the construct design and testing of the different construct versions are described in more detail in the original paper (Shkarina et al., 2022). The blue light stimulation triggers the rapid activation and oligomerization of Cry2olig, which in turn results in the proximity-induced activation of effector domains and subsequent processing of downstream substrates, culminating in cell death. While Cry2olig alone responds to the blue light within seconds (Taslimi et al., 2014), the timing of cell death induction is defined by the kinetics of effector activation as well as availability and efficiency of the processing and activation of downstream substrates (such as apoptotic executioner caspases, necroptotic effector GSDMD, or pyroptotic effector MLKL); the first morphological features of cell death can usually be detected within minutes after the beginning of illumination. Procedure A. Generation of stable cell lines expressing optogenetically activated cell death effectors (optoCDEs) 1. Lentiviral particle production a. Prepare stocks of purified lentiviral plasmids encoding optoCDE constructs. We recommend using endotoxin-free midi-or maxi-prep kits for plasmid purification. b. Twenty-four hours prior to transfection, plate HEK293T cells in a tissue culture-treated 6-well plate at 5 × 10 5 cells/well in 2 mL of fresh cell culture medium. c. For each well, prepare the transfection mix, as summarized in Table 1, in 1.5 mL tubes. Mix gently and incubate at room temperature for 15 min. Note: Over the course of virus production, the transfected cells might change the morphology, round up, and lift, as a consequence of viral particle release. g. After 36-48 h, collect the virus-containing supernatants from transfected HEK293T cells into 15 mL Falcon tubes. At this step, the medium from the wells transfected with the same construct can be combined and further processed together. h. Using a 5 or 10 mL syringe, gently pass the supernatants through a 0.45 μm syringe filter to remove cell debris and collect the filtrate into the clean 15 mL Falcon tube. Supplement the virus-containing supernatants with 20 μM HEPES (final concentration) to facilitate viral particle stability. i. At this stage, the virus-containing supernatants can be either used directly or stored at -80 °C for several months.

Cell transduction
a. Seed the appropriate number of cells to be transduced. i. For adherent cells: we recommend seeding 0.8 × 10 6 -1 × 10 6 of cells per well of a 6-well plate, 6-24 h before transduction. Cells should be properly attached and cover 60%-70% of growth surface. The optimal seeding density can be adjusted to accommodate for the cell size and differences in the growth kinetics between different cell lines. We recommend using 2-5 mL/well of virus-containing medium for 6-well plates, 2 mL/well for 12-well plates, and 1 mL/well for 24-well plates.
ii. For non-adherent cell lines: cells can be collected, centrifuged, and resuspended in the viruscontaining medium immediately prior to transduction. The second centrifugation step (spinfection) can be performed either in plates or in 50 mL Falcon tubes (in this case, the cells need to be resuspended and transferred to the plates or flasks following the spinfection).
iii. Include an additional well of mock-treated (polybrene-only treated) cells to control for cell viability and transduction efficiency. b. Replace the culture medium with the virus-containing medium supplemented with 5 μg/mL polybrene to facilitate viral particle adhesion. c. To facilitate the infection, centrifuge the cells with the virus at 3,000× g for 1 h at 37 °C. For the plate centrifugation, seal the sides of the plates with tape to prevent the accidental spillover of the supernatants during plate transfer and centrifugation. Suspension cells can also be centrifuged in 50 mL Falcon tubes. After the completion of this step, move the cells to the cell culture incubator. d. After 6-12 h, aspirate the virus medium, wash cells 1-2 times with the pre-warmed DPBS, and add the appropriate volume of fresh growth medium. If cell density is too high, cells can be transferred into the larger vessel at this step. e. Let the cells recover for 24-48 h to increase the cell number and allow for the proper expression of the selection markers and/or antibiotic resistance genes. f. To select successfully transduced cells, replace the medium with fresh medium containing the appropriate antibiotic concentration. i. Use the non-transduced wells as the positive control for selection efficiency.
ii. The optimal antibiotic concentration for each cell line can vary and has to be determined using an antibiotic titration.
iii. For most cell lines, we have successfully used the selection medium containing 1-5 μg/mL of puromycin or 50-100 μg/mL of hygromycin B Gold. g. After 3-5 days, remove the antibiotic-containing medium and expand the surviving cells.

Cell line validation
a. To confirm the transduction efficiency, induce optoCDE expression by incubating the cells with 1 μg/mL doxycycline for 24-48 h (at this stage, cells need to be protected from light to prevent the optoCDE activation and cell death). Include the non-treated control (without doxycycline) to control for the leaky expression. b. The percentage of mCherry-positive cells can be quantified using fluorescent microscopy or FACS.
The efficiency of cell death induction can then be tested as described in sections B and C. Note: OptoCDEs are fused with mCherry allowing easy assessment of construct expression. c. Following selection and validation, the stable lines can either be frozen away in liquid nitrogen for storage or directly used for experiments.

B. Optogenetic induction and visualization of pyroptosis, necroptosis, and apoptosis using confocal microscopy
1. Cell preparation for imaging a. One day before the experiment, seed the cells into the 8-well tissue culture-treated μ-slides at the concentration of 0.5 × 10 5 -1 × 10 5 cells/well. i. If using other type of cell culture plates, the cell concentration should be adjusted to achieve 30%-70% of the density at the day of experiment. We recommend using a lower cell density for experiments focused on the high-resolution visualization of the dynamic cellular events during different types of cell death in adherent cells.
ii. For poorly adherent cell types (such as HEK293T, MCF7, or equivalent), the slides can be additionally pre-coated with 5% collagen. 1) Prepare the appropriate volume of 5%-10% dilution of bovine skin collagen in sterile PBS. Mix thoroughly by inverting. Avoid vortexing, as this may lead to the collagen precipitation. expression levels but can also result in increased cytotoxicity. c. To protect the cells from light following expression induction, construct a dark chamber by fully covering a petri dish with foil ( Figure 2). Alternatively, cells can be placed in a plastic box or an alternative type of light-impermeable container, which allows air circulation and humidification.

Figure 2. Examples of the chambers that can be used to protect optogenetically activated cell death effectors (optoCDE)-expressing cells from light
d. On the day of experiment: pre-heat the microscope to 37 °C. Ensure that the temperature is stabilized before the start of the imaging. e. Gently wash the cells with pre-warmed PBS and replace the cell culture medium with pre-warmed Opti-MEM or alternative imaging medium. At this stage, all the manipulations need to be performed under dim light or red-light conditions to avoid spontaneous optoCDE construct activation. f. Optional: for cell death detection, the following reagents can be directly added to imaging medium: i. Visualization of membrane permeabilization: 1) CellTox Green: 1:10,000 2) DRAQ7: 1:1,000 ii. Visualization of PS exposure 1) Annexin V Pacific Blue: 1:500 2) Annexin V FITC 1:1,000 3) Annexin V Alexa Fluor 647 iii. Apoptotic caspase activation 1) CellEvent caspase-3/7 green-it is recommended to pre-load the cells with the dye for 1 h before an experiment.

Microscope setup
a. Both 488 and 496 nm lasers can be used for Cry2olig activation. During continuous whole-field photoactivation experiments, Cry2olig stimulation can be coupled with simultaneous imaging of fluorophores in the green channel [in this case, it is preferable to set up this channel as the last in an acquisition sequence to obtain the non-stimulated (t = 0) images]. However, this is not possible when using pulsed activation and/or single-cell targeting; in this case, only red or far-red fluorescent dyes and proteins should be used for the visualization of cell death and other processes of interest. b. Set up the time-lapse imaging. Keep in mind that Cry2olig activation will cumulatively depend on both the laser power and the frame rate, so both need to be adjusted accordingly (e.g., if using higher frame rate, it is recommended to decrease the 488 nm laser power, while longer frame intervals might require increased laser intensity for Cry2olig activation). As an example, Figure 3 shows the relationship between the blue laser intensity and the percentage of pyroptotic cells detected after 30 min of illumination.  c. Acquire the time-lapse series. An example of the typical pyroptotic, necroptotic, and apoptotic cell morphology is shown in Figure 4. Note that the kinetics and morphological characteristics of each type of cell death vary among cell types and might also depend on the optoCDE construct expression level. ii. The optimal laser intensity and number of scanning iterations varies depending on construct expression level, cell type, and type of cell death, and has to be determined empirically.
iii. If the signs of cell death (such as membrane blebbing, DRAQ7, or Annexin V positivity) are observed in the neighboring cells, reduce the ROI size and/or laser intensity or number of scanning iterations. b. Set up the time-lapse experiment. This should include several frames before the photoactivation as the baseline, after which the ROI stimulation is performed, and additionally 30-60 min or more after. c. Acquire the time lapse. To monitor for cell death, the imaging medium can be supplemented with the far-red cell death dyes (Annexin V or DRAQ7). The example of such experiment is shown at Figure  5. a. Set up the ROI stimulation experiment as described above. In this experiment, the ROI can include the whole field of view or be limited to the single cell. b. Reduce the 488 nm laser power to 0.1%-0.2% and number of iterations to 1-3. c. To monitor membrane permeabilization, supplement the imaging medium with DRAQ7. Use higher dye concentration (1:100-1:500) and laser power/gain at this stage to detect low amount of membrane damage. d. Perform the imaging and photoactivation. The transient low-level optoCDE activation will likely lead to three phenotypes: 1) the cells that will undergo cell death (acquire "high" DRAQ7 staining); 2) the survivor cells (might acquire moderate DRAQ7 staining and transiently display some early features of cell death, such as membrane blebbing or nuclear condensation, but are able to revert to the normal morphology after), and 3) the cells that are not affected by the stimulation (not gaining DRAQ7 signal and no change in morphology) ( Figure 6). The ratio of these cells in the population might depend on the optoCDE expression level, as well as additional cell-intrinsic factors (such as the variability in expression of the downstream effectors or activity of membrane repair systems). e. Adjust the stimulation parameters to achieve the desired optoCDE activation and cell survival ratio and repeat the experiment.

C. Optogenetic induction of pyroptosis in human macrophage-like cell lines
Important points 1. In this protocol, we utilize U937, human monocyte-like cell line, which can be differentiated into the macrophage-like phenotype using PMA treatment. However, a similar type of experiments can be performed with other cell lines. 2. Prior to seeding, transgenic U937 lines are grown in suspension in T75 flasks in complete RPMI medium.
For optimal growth, the medium is exchanged every 2-3 days. Avoid growing the cells to too high density, as this will lead to the reduction of cell viability and might impact the differentiation.
Step-by-step protocol 1. Cell seeding and differentiation a. Collect the desired volume of U937 cells into the 50 mL flasks and centrifuge at 300× g at room temperature for 5 min. Discard the medium. b. Resuspend the cells in fresh RPMI. c. Count the cells and resuspend to the final density of 250,000 cells/mL. d. Add PMA to the final concentration of 5 ng/mL. e. Seed in 24-well black tissue culture-treated plates (4titude). Include additional wells for the nonilluminated cells and the positive (total lysis) control. f. After 24 h, remove the PMA-containing medium, wash once with pre-warmed PBS, and add fresh growth medium. g. After 48 h, replace the growth medium with the induction medium containing 2 μg/mL doxycycline and incubate cells overnight to induce optoCDE expression.

Illumination
a. Set up the light plate apparatus in the cell incubator ( Figure 6B). If using transient illumination times, these experiments can be performed on the bench before moving the plates to 37 °C. b. Generate the custom illumination program using IRIS software. The example of an illumination layout is shown in Figure 7C. We recommend using duplicate wells for each illumination condition and including additional wells for the non-illuminated controls and total cell lysis (cells treated with 1% Triton X-100). c. Download the program and transfer the files to the micro-SD card. Perform a short test illumination round to ensure that the device works properly. d. Remove the cell culture medium from the cells and add 300 μL of pre-warmed Opti-MEM or equivalent serum-free phenol red free medium per well. e. Place the plate into the light plate apparatus, cover the lid, and start the illumination. Upon the end of the program, the lights should turn off automatically. f. Remove the plate from the light plate apparatus. Add 30 μL of 10% Triton X-100 to the positive control (total cell lysis) wells. Using a P1000 pipette, pipette 5-10 times to lyse the cells. g. Centrifuge the plate at 500× g for 5 min at room temperature to pellet the cell debris. h. Gently collect the supernatant from each well to the pre-labeled 1.5 mL microtubes. Avoid touching the well bottom to prevent debris collection. At this stage, 100 μL of supernatant can be used for lactate dehydrogenase (LDH) measurement, while the rest can be preserved at -20 °C for the cytokine measurement. 3. Assessment of cell lysis using LDH activity assay a. Prepare the appropriate volume of the LDH reaction mix. b. Using a multichannel pipette, add 30 μL of reaction mix per well of a flat-bottom 96-well plate. Avoid creating bubbles, as this will impact the assay efficiency and reading. c. In each well, add 30 μL of the cell supernatants collected in the step 2h. Include two to three technical replicates per each experimental well. Minimize delays while pipetting, as this might impact the difference in the colorimetric reaction between the wells and lead to bias. d. Mix by gently tapping the plate on the side. Avoid mixing by pipetting, as this will generate bubbles.
If bubbles are produced during pipetting, they can be manually removed before reading the plates using either syringe needles or an inverted Bunsen burner. e. Incubate the plate for 15-20 min at room temperature, protected from light. f. Add 30 μL of stop solution per well and gently mix. g. Measure the absorbance using a 490-492 nm filter. 4. Assessment of the IL-1β release a. IL-1β release quantification following illumination is typically quantified using ELISA, performed according to the manufacturer's protocol (available at R&D website). b. We recommend using several dilutions (1:1, 1:2, 1:5) of the collected supernatants, as the amount of IL-1β in undiluted supernatants frequently exceeds the dynamic range of assay. c. Alternatively, a FRET-based no-wash homogeneous time resolved fluorescence assay can also be utilized for quicker single-step IL-1β quantification.

Western blot analysis of optoCDE activation and cell death
a. Before the start of the experiment, prepare the following reagents: i. Pre-labeled 1.5 mL microtubes for supernatant and lysate collection.
ii. Cell lysis (RIPA or equivalent) and loading buffer.
iii. Reagents for phenol-chloroform protein precipitation. iv. Reagents for the polyacrylamide gel preparation (or pre-cast gels). v. Western blot running, transfer, and wash buffers. b. Perform the illumination of the plates as described in step 5a. Ensure that the cells are in the good condition before the start of experiment and are mCherry-positive (this can be assessed using epifluorescent microscopy). c. Centrifuge the plates at 300-500× g for 5 min at room temperature to pellet the dead cells and collect the supernatants from each well into 1.5 mL microcentrifuge tubes. d. Immediately add 50-70 μL of hot (95 °C) loading buffer to the remaining adherent/pelleted cells. At this stage, plates can be processed immediately or sealed with parafilm and stored at -80 °C for later processing.

13
Published: Jul 20, 2023 e. Pipette the lysis buffer up and down 5-10 times and collect the lysates into the microtubes. If lysates become too viscous at this stage, they can be additionally sonicated. f. Incubate for 5 min at 95 °C. g. In parallel, perform the protein precipitation in the cell supernatants using the phenol-chloroform protein extraction method (Demarco et al. 2022). After the precipitation, the dried protein pellets can be either resuspended in 1× lysis/loading buffer and processed independently or combined with lysates of corresponding wells. h. Load samples on the 10%-12% polyacrylamide gels. i. Perform the gel running and western blot analysis of the samples following the protocol available in the host lab. The detailed protocol for western blot analysis of such samples can also be found in Demarco et al. (2022). j. For the detection of inflammatory opto-caspase activation, we recommend using the following primary antibodies: rabbit anti-cleaved IL-1β (83186, CST; 1:1,000), which detects IL-1β by activated opto-(h)caspase-1; mouse anti-IL-1β (12242, CST; 1:1,000), which detects full-length unprocessed IL-1β in non-activated cells; rabbit anti-GSDMD (ab210070; 1:1,000; Abcam), which detects nonactivated GSDMD in resting cells; rabbit anti-cleaved N-terminal GSDMD (ab215203; 1:1,000; Abcam), which detects active GSDMD cleaved by opto-(h)caspase-1; mouse anti-caspase-1 (clone Bally-1 AG-20B-0048-C100; 1:1,000; AdipoGen); mouse anti-mCherry (ab125096; 1:2,000; Abcam); and HRP-conjugated mouse anti-tubulin (ab40742; 1:5,000; Abcam), used as a loading control. The secondary HRP-conjugated or fluorescently labeled antibodies can be used according to the lab's choice. A representative blot is shown in Figure 8.

Analysis of microscopy images
The analysis of the microscopy images is described in the Methods section of the original publication (Shkarina, et al., 2022). For quantification of different types of cell death, we assessed several parameters: a) appearance of the characteristic morphological changes associated with the each type of cell death (cell rounding and swelling, nuclear condensation for pyroptosis and necroptosis, nuclear fragmentation and persistent membrane blebbing for apoptosis); b) Annexin V staining, as an indicator of membrane scrambling and phosphatidylserine exposure (for all three types of cell death); and c) uptake of specific dyes associated with the loss of membrane integrity, such as CellTox Green or DRAQ7 (for pyroptosis and necroptosis). Additionally, activation of the apoptotic caspases can be monitored using genetically encoded or chemical fluorogenic caspase reporters (such as VC3AI, described in the original paper, or CellEvent caspase-3/7 reporter system). The number of dying cells can be normalized to the total number of cells per field of view, or, alternatively, to the number of mCherrypositive cells in the population, to account for the construct expression and transduction efficiency.

Analysis of LDH assay and ELISA results
LDH release from pyroptotic (or generally necrotic) cells following illumination is quantified using the following equation: (LDHsample -LDHnegative control)/(LDHpositive control -LDHnegative control) × 100 Where negative control is assay medium (e.g., Opti-MEM or other phenol red free medium) and positive control corresponds to the cells lysed with the 1% Triton X-100. Quantification of IL-1β release is performed according to the manufacturer's protocol. Note that the variation in initial (pre-illumination) cell density between the cell lines or conditions can have a strong effect on the amount of detected IL-1β. This can be corrected by normalizing IL-1β values to the ratio of maximum LDH values from lysed cells. An example of the LDH and IL-1β secretion data obtained from this type of experiments is shown in Figure 9.